Bacterial cellulose. Cellulose is the most abundant biopolymer on earth, synthesized by plants, fungi, algae, bacteria and some animals. Its chemical structure is a polymer composed of glucose monomers linked thru β-1,4. Most celluloses occur as a crystalline allomorph cellulose I, the native allomorph, which can be treated to form into other allomorphs such as cellulose II, III, IV synthetic products. Cellulose I is composed of both Iα and Iβ crystalline unit cells. Cellulose Iα is triclinic with dimensions a=6.3 Å, b=6.9 Å, c=10.36 Å, α=113.0°, β=121.1°, γ=76.0° (Vietor, 2000). Cellulose Iβ is monoclinic with dimensions a=8.17 Å, b=7.86 Å, c=10.38 Å and γ=97.0° (Brown, 1984). Morphologically, cellulose I exists as submicroscopic rods known as microfibrils, the shape and size of which vary and are governed by the genetics of the organisms that generate it. The most common source of cellulose is from higher plants such as cotton and ramie producing microfibril widths of 2-5 nm. Algal, bacterial and tunicate celluloses forms larger microfibrils of 15-30 nm in width and these celluloses are rich in Iα crystalline.
Bacterial cellulose is a product of microbial primary metabolism. For example, cellulose is produced by species such as Zoogloea, Sarcina, such as Sarcina ventricula (Canale-Parola1960), Salmonella, Rhizobium (Napoli, 1975), Pseudomonas, such as Pseudomonas fluorescens (Spiers, 2003), Escherichia, Agrobacterium, such Agrobacterium tumefaciens (Matthysse, 1995), Aerobacter, Achromobacter, Azotobacter, Alcaligenes, and Acetobacter, also known as Gluconacetobacter. The most studied and used cellulose-producing bacteria specie is Acetobacter xylinum, which includes the strains ATCC 23769, 10145, 53582, AX5 and many others (Brown, 1996) (Klemm, 20010. Microorganisms of Acetobacter are obligate aerobes and generally are found in fruits, in vegetables, most likely in rotting fruits and vegetables, in vinegar, fruit juices and alcoholic beverages (Klemm, 2001).
When a liquid medium, known as H-S medium (Hestrin, 1954) that consists of 2 wt % D-glucose, 0.5 wt % peptone, 0.5 wt % yeast extract, 0.27 wt % disodium phosphate, 0.115 wt % citric acid (monohydrate), and distilled water is inoculated with a strain of Acetobacter xylinum, a cellulose pellicle will be formed on the air-liquid medium interface. Glucose functions as bacteria's carbon source, peptone as nitrogen source, yeast extract as vitamin source and citric acid and disodium phosphate as buffer system for the medium. Before the medium is inoculated with a bacteria strain, it goes through sterilization by autoclaving. During this sterilization process, D-glucose is partially isomerized to D-fructose thus degrading to dark-yellow products resulting into a yellow liquid growth medium. About 6% of D-glucose will be lost due to transformation to fructose. To stabilize D-glucose and to minimize its loss, citric acid, which is part of the buffer system acts as a stabilizing agent. The mechanism of cellulose formation by Acetobacter xylinum is as follows; the bacteria increase their population by consuming glucose and oxygen initially dissolved in the liquid medium, when the oxygen has diminished, only bacteria having access to air can continue cellulose-producing activity, thus forming the cellulose pellicle at/in the air-liquid medium interface. The bacteria below the surface area are considered dormant but can be reactivated by using the liquid as an inoculum for a new culture medium. A. xylinum cells when fed with glucose cause a slow evolution of carbon dioxide as it forms cellulose. The gas accumulates on the surface of cellulose fibrils and is believed to be another cause of the cellulose pellicle flotation on the air-liquid interface. The increase of mass or thickness of cellulose pellicle occurs at the upper part of the pellicle surface, where oxygen is accessible. The cellulose polymers diffuse through the cellulose pellicle sheet to access the oxygen. The oldest part of the pellicle is the under part, which has been pushed progressively downwards into zones of decreasing oxygen pressure and lesser activity. The monosaccharide is converted by Acetobacter xylinum dehydrogenase into (keto)gluconic acids. D-glucose not only acts as a carbon source but also as a cellulose precursor.
Bacterial cellulose is potentially useful in many applications, but large-scale production of this material is yet to be developed. Static culture, which is a better production method requires wide (e.g. extended, substantial) surface area since the cellulose pellicles form at/in the air-liquid surface. However, highly extended (e.g., wide) culture surface areas are impractical and thus unsuitable for large-scale cultivation, and further improvements are needed to provide economical processes for bacterial cellulose production.
Numerous attempts have been made to increase production of bacterial cellulose, but none have yet proven feasible. Vandamme (Vandamme, 1998) and coworkers, for example ascertained that improvement of bacterial cellulose production could be achieved by proper strain selection, mutation, medium composition optimization, and physico-chemical fermentation parameter control. These authors combined nutritional, genetic and bioprocess-technological optimization in attempts to obtain high levels of cellulose production, and with agitated culture demonstrated that cellulose formation is enhanced by adding insoluble microparticles such diatomaceous earth, silica, small glass beads and loam particles. Another way of enhancing production in agitated culture is by adding ethanol, since ethanol is metabolized during the growth cycle of Acetobacter, producing more cellulose as transformation of glucose to fructose is inhibited. Jonas and coworkers' (Jonas, 1998) extend the Vandamme methods by using pH control to improve production, finding an optimal pH range of 4 to 7. One method of pH control, for example, is by using HCl and NaOH as described by Hestrin and Schramm (Hestrin, 1954). There is also an optimal temperature for cellulose production by Acetobacter strains, the range lying between 20° C. to 30° C. Most authors use 28-30° C. (Vandamme, 1998). In 1963, Webb and Colvin (Webb, 1963) added plant extracts from a number of plant sources resulting in increased cellulose synthesis by A. xylinum. The plant extracts were from tomatoes, carrots, potatoes, oranges and spinach leaves. Addition of endo-1,4-glucanase from Bacillus subtilis also enhanced bacterial cellulose production. When glucanase was added, the structural properties of the bacterial cellulose were not modified (Tanouchi, 1995). Joseph (Joseph, 2003) used polyacrylamide-co-acrylic acid to enhance bacterial cellulose production in shake culture, but the morphology of the product was changed when compared to the native product. Static culture is the most common method to grow bacterial cellulose. Aside from shaking or agitating cultures, bacterial cellulose can also be produced using an airlift reactor equipped with draft tube and riser; or using a rotating disk reactor where part of the surface of the disk is alternately positioned between the liquid medium and the atmosphere (Shoda, 2005).
Bacterial cellulose is primarily composed of Iα crystalline forms with relatively small amounts of Iβ present. Acetobacter cellulose, for example, is estimated to consist of about 60% to 70% Iα, which is very different from cotton cellulose that comprises 60% to 70% Iβ (Atalla, 1984). The presence of stress during the development of cellulose is believed to cause the formation of the Iα crystalline form. In crystallization of Japanese cypress tracheid cellulose, the stress is frequently exerted by the growing cells which stretch the primary cell wall of the plant examined. Iβ phase forms when the stress is relieved due to the fluidity of the environment thereby result in the presence of both Iα and Iβ phases in the tracheid cellulose (Kataoka, 1999). The ratio of Iα and Iβ can be determined using FT-IR spectra, where a peak at 750 cm-1 represents Iα, and a peak at 710 cm-1 represents Iβ (Id).
Physical properties of bacterial cellulose can be modified in many ways. When hemicellulose-like saccharides are added into the liquid medium growth of bacterial cellulose, its microfibril aggregation patterns are modified, whereby Iα-type crystalline arrangements transform to Iβ crystalline type. Thus, hemicellulose addition appears to transform bacterial cellulose to the higher plant cellulose structure (Uhlin, 1995). A drying process can alter the degree of crystallinity of cellulose but will not change Iα/Iβ ratio (Udhardt, 2005). Isolation procedures may also have an effect on the structure of bacterial cellulose (Uhlin 1995).
Bacterial cellulose is an excellent alternative to plant cellulose, particularly in areas where plant cellulose can't be used, such as where a high purity crystalline cellulose structure is essential. Table 1, for example, lists some of the patented cellulose products and applications (e.g., in the health care sector where its biocompatibility is recognized). ‘Never-dried’ bacterial cellulose has been demonstrated to have suitable biocompatibility for use in wound healing (Czaja, 2006). Additional known uses include stereo headphone diaphragms, food, paper, chromatographic techniques, cosmetics stabilizer and for latex binders. Another application is in membrane technology as BC has highly porous feature. The major utilization of BC is in biomedical or health sector. The unique nanostructure of bacterial cellulose gives it a high mechanical strength and remarkable physical properties in wet and dry state thus making this material very functional in many applications. However, although significant applications have been realized, mass production has not been viable.
TABLE 1Art recognized cellulose products and applications thereof.Cellulose ProductUses and ApplicationTemporary artificial skinTreatment for burns, ulcers(Biofill ®, Bioprocess ®,and dental implantsGengiflex ® (Biofill01)[Biofill02]Nonwoven paper or fabricEnhance the property of latex(Weyerhaeuser),or binders, repair old documents[Biopolymer]Diaphragms (Cannon, 1991)Stereo headphonesMicrobial celluloseImmobilization of proteins,(MC_patent)chromatographic techniquesFood, food or diet fibersubstituteStabilizer, viscosity modifiersBASYC (Klemm, 2001)Artificial blood vessels formicrosurgery Protective cover formicronerve structureMembranes (Choi, 2004)Environmentally compatibleion-exchange membrane
Nanocomposites. Nanocomposites are a new class of composites characterized by ultra fine phase dimensions of 1 nm to 1000 nm. Such nanocomposites include hybrid materials comprising polymer matrix reinforced with a nanoscale reinforcement (e.g., fibers or platelets). There are three main classifications of nanocomposites reinforcements: (i) nanoscale level in three dimensions such as spherical silica; (ii) elongated reinforcements with nanoscale level in two dimensions such as fibers and carbon nanotubes (iii) sheet-like structures with nanoscale level in one dimension such as layered silicates, mica and clay.
The outstanding properties of bacterial cellulose that make it excellent as a reinforcing material for nanocomposites include but are not limited to: high purity, without the blend of lignin and other hemicellulose as with plant; high crystallinity; biodegrability; water holding capacity up to 100-times its weight; and excellent biological affinity. The reinforcing effect of cellulose comes from its ‘whiskers’ percolating network and good interfacial compatibility with the polymer matrix. When the cellulose and its polymer matrix do not have a good interaction or good miscibility, the nanocomposite is likely to possess inefficient mechanical and physical properties. Interactions of the polymer matrix with cellulose are closely associated with the following factors: solubility of polymer additives, diffusibility of the additives to each microfibril surface and the extent of hydrogen bonding between the additives and microfibrils. Designation of a polymer for cellulose nanocomposite involves determination or estimation of the interfacial compatibility between polymers based qualitative considerations, and thus description of precise atomistic scale interfacial phenomena is difficult to assess. Currently, the choice of the polymer matrix is typically based primarily on trial and error. One of the properties of a material that can give an approximation of its behavior when interacting with another material to form into a nanocomposite is the solubility parameter (δ). Addition of polymers that have solubility parameters close to cellulose can affect cellulose aggregation. There are numerous existing approaches of determining the solubility parameter (δ). For example, one approach is to use functional group contributions of a material or a polymer (VanKrevelen, 1976). The solubility parameter for cellulose was evaluated by Bochek (Bochek, 1993), who obtained 56.2 (J/cm3)1/2. When compared to the δ values calculated using other methods, Bochek's evaluation is greater, based on taking into account the highly polar characteristic of cellulose, where the polar characteristics are the number of hydrogen bonds and their energies. The values of cellulose δ acquired from other methods are (method: δ): Small: 21.0 (Small, 1953), Hoy: 29.6 (Hoy, 1970), Van-Krevelen: 38.8 (VanKrevelen, 1976), Fedors: 34.9 (Fedors, 1974), all δ are in units (J/cm3)1/2.
Acetobacter synthesized bacterial cellulose has been grown in static culture the presence of limited amounts of particular polymers to study details of microfibril formation (aggregation) and crystallization, or understand the functionality of cellulose in plants. The polymers used include hemicellulose (Uhlin, 1995) (Yamamoto, 1994) (Haigler, 1982), Calcofluor (Haigler, 1980), Calcofluor White or Congo Red dyes (Colvin, 1983), xylan (Yamamoto, 1996) (Yamamoto, 1994) (Ohad, 1963), phosphomannan (Ohad, 1963), xyloglucan (Whitney, 19990 (astley 2003), pectin (Astley, 2003) (Ohad, 1963), glucomannan and galactomannan (Whitney, 1998), etc. The aggregation of bacterial-produced subfibrils has been shown to be altered in the presence of particular agents. For example, bacterial cellulose synthesized in the presence of the fluorescent brightener Calcofluor under static pellicle culture conditions (Haigler, 1980), indicates that Calcofluor prevents the assembly of crystalline cellulose microfibrils and ribbons by Acetobacter zylinum, where Calcofluor alters cellulose crystallization by hydrogen bonding with glucan chains. Bacterial cellulose has been synthesized in the presence of hemicellulose under static pellicle culture conditions (Uhlin, 1995, Yamamoto, 1994, Haigler, 1982), where patterns of aggregation of the bacterial cellulose were modified to be more like the plant Iβ-type than the bacterial Iα-type when the cellulose was produced in the presence of hemicellulose-like saccharides (xyloglucan; mannan, xylan and carboxymethyl celluose). Other polymers studied include: dyes (Colvin, 1983), xylan (Yamamoto, 1996) (Yamamoto, 1994) (Ohad, 1963); phosphomannan (Ohad, 1963), xyloglucan (whitney 1999) (Astley 2003); pectin (Astley 2003) (Ohad 1963); and glucomannan and galactomannan were added to the growth medium of BC-producing bacterium, forming thinner diameter cellulose ribbons (Whitney, 1998). These studies, however teach nothing about the distribution or properties of cellulose fibrils compositions comprising such particular polymers.